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Mouse models are a valuable tool for studying acute injury and chronic remodeling of the myocardium in vivo. With the advent of genetic modifications to the whole organism or the myocardium and an array of biological and/or synthetic materials, there is great potential for any combination of these to assuage the extent of acute ischemic injury and impede the onset of heart failure pursuant to myocardial remodeling.

Here we present the methods and materials used to reliably perform this microsurgery and the modifications involved for temporary (with reperfusion) or permanent coronary artery occlusion studies as well as intramyocardial injections. The effects on the heart that can be seen during the procedure and at the termination of the experiment in addition to histological evaluation will verify efficacy.

Briefly, surgical preparation involves anesthetizing the mice, removing the fur on the chest, and then disinfecting the surgical area. Intratracheal intubation is achieved by transesophageal illumination using a fiber optic light. The tubing is then connected to a ventilator. An incision made on the chest exposes the pectoral muscles which will be cut to view the ribs. For ischemia/reperfusion studies, a 1 cm piece of PE tubing placed over the heart is used to tie the ligature to so that occlusion/reperfusion can be customized. For intramyocardial injections, a Hamilton syringe with sterile 30gauge beveled needle is used. When the myocardial manipulations are complete, the rib cage, the pectoral muscles, and the skin are closed sequentially. Line block analgesia is effected by 0.25% marcaine in sterile saline which is applied to muscle layer prior to closure of the skin. The mice are given a subcutaneous injection of saline and placed in a warming chamber until they are sternally recumbent. They are then returned to the vivarium and housed under standard conditions until the time of tissue collection. At the time of sacrifice, the mice are anesthetized, the heart is arrested in diastole with KCl or BDM, rinsed with saline, and immersed in fixative. Subsequently, routine procedures for processing, embedding, sectioning, and histological staining are performed.

Nonsurgical intubation of a mouse and the microsurgical manipulations described make this a technically challenging model to learn and achieve reproducibility. These procedures, combined with the difficulty in performing consistent manipulations of the ligature for timed occlusion(s) and reperfusion or intramyocardial injections, can also affect the survival rate so optimization and consistency are critical.

When the mice are unresponsive to toe-pinch, a sterile lubricant (Tears Renewed) is applied to the eyes to protect them from desiccation and the left side of the chest is coated with depilatory (e.g. Nair) to remove fur from the skin.

The depilatory is washed away with warm running water and betadine/alcohol swabbing is used to disinfect the surgical area.

The mouse is placed on a warm deltaphase isothermal pad which is fixed to a plexiglass table. Each limb is immobilized using tape and a thick thread is placed horizontally under the top teeth to hold the upper jaw in place.

The table is positioned vertically and a fiberoptic light is shone directly onto the neck region for transesophageal illumination. This requires precise placement such that the opening of the throat is viewed as a well lit orifice, thus enabling the trachea to be visualized to facilitate insertion of the PE tubing.

The tubing is then connected to the ventilator (connected to a 95% O2/5% CO2) to administer constant positive pressure ventilation (TOPO ventilator; rate 125 breaths/min; peak inspiratory pressure 10-12 cmH2O; *note: settings vary with strain and gender 1-3). Once ventilation is confirmed by synchronous chest movements, the connection is fixed to the pad with tape to avoid extubation during the surgery.

Using toothed forceps to pull skin up and away from the chest, a #10 sterile scalpel blade attached to a #3 scalpel handle is used to make a 1.5cm incision in the skin parallel to the sternum.

Curved Vanna microscissors are used to cut the pectoralis muscles and make a small hole in the intercostal muscle.

Straight, blunt microscissors are used to cut through 3 ribs.

A 9mm pediatric ophthalmic speculum is used to retract the rib cage.

Using the curved forceps, pull the pericardium away from the heart and use the toothed forceps to gently tear it open.

Using the Castroviejo needle holder, a 6mm tapered point 3/8 needle threads the 8-0 polyethylene suture underneath the left anterior descending coronary artery (along the long axis of the heart) perpendicular to it.

For a temporary ligature that can be removed for timed reperfusion, a sterile 0.5-1cm piece of PE90 is placed on the heart in parallel to the coronary artery. The suture, which has first been looped under the coronary artery, is then tied to the tubing. At the time it is to be released, the ligature is loosened. This can be repeated as desired and the time of occlusion/reocclusion can be modified 4. Depending on the length of the protocol and the type of anesthesia used, supplementation may be necessary.

For a permanent occlusion, the ligature laced under the coronary artery is simply tied. Blanching and dyskinesia are apparent and the long end of the suture is cut 5-10.

For intramyocardial injection(s), a sterile Hamilton syringe with a 30 gauge sterile beveled needle is introduced into the base of the heart above the area of injury on the right side of the ligature. The needle is then advanced into the area of injury and withdrawn slightly so that the bevel can be seen approximately at the border zone. Some of the solution in the syringe (2-3μl) is injected into the heart and the needle is held in place. The syringe is withdrawn another 1-3mm and the rest of the solution is injected. The syringe is held in place until the bleb that is formed by the solution dissipates. The needle is then removed. If there is any bleeding, a cotton-tipped applicator is gently pressed onto the needle insertion site until the bleeding stops 5-7.

Once the myocardial manipulations are complete, the rib retractors are removed and the thoracic cavity is closed with 2-3 mattress sutures using 6-0 surgipro suture.

Two-three mattress sutures are then made to close the pectoralis muscles, 1-3 drops of 0.25% marcaine 1:10 in sterile saline (0.1ml/25g mouse) is applied to the muscle and then 2-3 mattress sutures are made to close the skin.

The mouse is removed from the ventilator. Once rhythmic, rapid, shallow breathing is verified, the mouse can be extubated.

0.5ml warm sterile saline is injected into the dorsal subcutaneous space and the mouse is placed on a warming pad in a cage until it regains mobility (1 hour minimum).

For survival experiments, mice are placed back into their cages and returned to the vivarium until the time of sacrifice. During first 2 days, moistened food is placed on the cage floor to facilitate feeding (so they don't have to reach up which may cause pain) and buprenorphine should be administered every 6-12hr. Post-operative care also includes daily monitoring for the first week to verify adequate mobility, grooming, and eating habits.

The surgical instruments are wiped clean with ethanol and inserted into the bead sterilizer before the next surgery.

At the time of sacrifice, mice are anesthetized with sodium pentobarbital (65mg/ml; 55-65 mg/kg). When an adequate plane of anesthesia is achieved, the thoracic cavity is opened.

While the heart is still beating, a syringe with a 23 gauge needle containing cold potassium chloride (KCl, 30mM) or 2,3-butanedione monoxime (BDM; 10mM) is used to puncture the posterior basal region of the ventricle and the solution is slowly injected into the chamber until the heart is arrested in diastole.

Once the heart is removed, a syringe containing PBS is used to retrogradely perfuse rinse the heart to remove any blood that remains. For acute studies, at the end of the reperfusion period, the LAD is re-ligated at the original point of occlusion. A solution containing 1% Evan's blue is injected into the aorta. Once the heart is extracted, it is cut transversely into 3 sections of equal thickness, incubated in 1% 2,3,5-triphenyltetrazolium chloride, and imaged for morphometric analyses 11. For chronic studies, the heart is then immersed in fixative, then processed and embedded according to routine procedures. Slides can then be stained histologically and imaged for morphometric analyses (using Scion, NIH Image J, or Image Pro Plus) 9,10,12.

Representative Results:

When done correctly, the survival rates in mice (male: age 8-10 weeks, 22-28g; female: age 10-12 weeks, 20-26g) are: over 90% in acute ischemia/reperfusion and ischemic preconditioning experiments, over 85% in permanent artery ligation studies, and approximately 80% for intramyocardial injections. Since early injury is more readily visible by metabolic changes rather than structural, infarct size determination in ischemia/reperfusion and ischemic preconditioning experiments is performed by infusing 1% Evan's blue dye into the aorta which will perfuse the heart that is not supplied by the LAD (Figure 1A). Once the heart is removed and transversely cut in half, the tissue is incubated in 1% solution of 2,3,5-triphenyltetrazolium chloride to measure infarct size (Figure 1B). The areas are measured using Scion or NIH imaging software which can be calibrated using a micrometer imaged at the same magnification These numbers are used to calculate area at risk/left ventricle and infarct size/area at risk 11. Strain differences can result in variations in body weight and heart size and so care should be taken to normalize these measures to heart weight, body weight, or tibia length for comparative purposes.

Permanent artery ligation results in gross structural changes such as necrosis, wall thinning, and chamber dilation. Comparison of the effects of treatment and/or time on infarct size and necrosis relative to the left ventricle, chamber area, septal wall and left ventricular free wall thickness in the permanent occlusion model (Figure 2A) can also be measured using Scion or NIH imaging software. Collagen staining with picrosirius red/fast green (Figure 2B) can be used to measure insterstitial fibrosis which correlates to functional indices of wall stiffening8-10. The image in Figure 3 represents the distribution of 6ul solution (Evan's blue) injected into the border zone of the heart following permanent artery ligation. Notice that it proceeds in the direction of the injury as well as toward the base and also transmurally.

Figure 1. A. Evan's Blue injected into the aorta prior to excision. This image shows the perfused regions of the heart (stained) and the occluded area (unstained). B. Evan's Blue and TTC staining following acute ischemia/reperfusion injury. This is a representative image (20x) showing the blue dye distribution which stained the unoccluded regions as well as TTC staining of metabolically viable tissue (red). Necrotic areas do not stain and so they remain pale (outlined).

Figure 2. A. Hematoxylin and eosin stain. This is a representative image (20x) of H&E staining of a mouse heart cut transversely through the infarct region at 4 days post-MI (20x). The * denotes tissue necrosis, arrows point to granulation tissue, RV = right ventricle and LV = left ventricle. B. Picrosirius red and fast green stain. This is a representative image (20x) of picrosirius red/fast green staining of a cross-section of mouse heart at 4 weeks post-MI. The cytoplasm stains green and collagen fibers are red.

Figure 3. Evan's Blue dye stain distribution following 6ul intramyocardial injection. This is a representative image showing the global and transmural distribution of Evan's Blue dye throughout the heart following 6ul intramyocardial injection at the border zone immediately following coronary artery ligation (12x).

Coronary heart disease continues to be an epidemiologically and fiscally significant public health problem. Considerable basic research is still needed to understand the mechanisms by which injury and remodeling proceed and how potential therapeutics may modulate these processes if they are to be developed for clinical use. Rodents are most commonly used and the wide range of genetically modified mice available makes this species a more attractive model.

Although there are differences between mice and other species, there are many advantages to a murine model. Use of a simple dissecting scope or magnifying glass and well lit conditions enable the vasculature to readily be seen (for detailed gross anatomy of vasculature, see Salto-Tellez et al., 200413). To reduce the risk of post-operative mortality, it is very important to avoid severing large vessels since the total blood volume of a 25g mouse is less than 2ml14. In the event that excessive bleeding occurs, gentle application of pressure or pinpoint cauterization can be used to stop the bleeding.

This procedure can also be modified in a variety of ways. For example, mice can be anesthetized using isoflurane, ketamine/xylazine, or sodium pentobarbital and appropriate selection is determined by the duration of the protocol15-18. The toe-pinch reflex is the most commonly used index of the depth of anesthesia. Further, to improve the probability for long term survival, some investigators use antiarrhythmic drugs such as lidocaine to reduce the incidence of lethal arrhythmias19,20 however, it must be taken into account that this has recently been shown to have antiapoptotic properties in an acute model21. Also, to reduce post-operative pain, analgesics such as buprenorphine can be administered for the first 48 hours after surgery3,16,17,22,23. To maintain body temperature during surgery (especially for longer protocols), a rectal probe in series with a heating pad is often used in place of the isothermal pad. For ischemia/reperfusion and/or ischemic pre- or postconditioning: the duration of the occlusion(s) and reperfusion(s) can be altered; for permanent occlusion, the size of the infarct may be modified by adjusting the location of the ligature; and for intramyocardial injections (eg. cells, proteins), there can be 1-3 injection locations and the volume per injection can be up to 15 μl24. If cells are being injected, the gauge of the needle used (usually 26-30)5,25,26 should be chosen based on the size of the cells so the internal diameter of the needle is large enough to avoid sheering. To avoid confounds due to inflammatory processes triggered by the surgery, some investigators have reported using a snare that is manipulated ex vivo to occlude and reperfuse the hearts in a closed chest mouse at any point after the surgery27-29. More recently, Gao et al.30 have shown that temporary and permanent occlusion can be performed without the need for ventilation and a few laboratories have begun to use ultrasound to perform closed-chest intramyocardial injections25,31.

Since the first study demonstrating feasibility of ligating the coronary artery in mice was published by Johns and Olson in 195432, many others have adopted this model and modified it to study various aspects of myocardial injury and remodeling3,33-45. The nature of mice in terms of size, reproductive capacity, and comparatively less expense for purchase and maintenance make this species an appealing tool for a broad range of physiologic and pathophysiologic studies. As the miniaturization of technology for imaging in vivo advances46-49, as well as means to perform and analyze large scale genomics and proteomics, drug screening, efficacy of cell-based and/or protein therapies as well as biomaterials50-64, combined with the increasingly wide range of genetic manipulations afforded by ubiquitous or tissue specific transgenic or mutant/knockout mice, the murine model of myocardial infarction will undoubtedly continue to be an invaluable tool in evaluating acute cardiac injury and long term remodeling. Therefore, there is unquestionable value in being able to perform these experiments reliably and reproducibly.

I would like to acknowledge the Department of Research and Graduate Studies for providing funds to support my research and the Department of Comparative Medicine for their vigilance and assistance. I would also like to recognize the Department of Physiology for their support and guidance as well as the students and technicians in my lab for their help. Lastly, I would like to thank my post-doctoral mentor, Dr. Charles E. Murry, for the training opportunity during which time I learned the mouse microsurgery.

15 Comments

In reviewing this and a couple other LAD models, I note that all use 8-0 Prolene mono-filament, or alternative polypropylene suture. With such a price difference, I am temped to use 8-0 nylon mono-filament. What are the pros and cons of prolene versus nylon? Is SURGIPRO II/SURGIPRO Nonabsorbable Monofilament Polypropylene from Suture Express less expensive than prolene? We have expire 7-0 prolene on Taper will that suffice if prolene is a must?

Preferences are operated-dependent. If you can use 7-0 or nylon without increasing drag and risk of tear, then I don't see any reason not to use it. I get a lot of mileage from a box of the VP-900-X and so the price gets distributed. Depends on how many you get in a box, ² needles or one, and how long each strand is.

I have recently started performing the MI model on mice using the TOPO ventilator. I have had several animals suffer non-recoverable pulmonary edemas, suggesting my inspiratory pressure is to high. Would you mind telling me the exact settings you set each dial to in order to achieve a peek inspiratory pressure of 10-1² cmH²O, and whether you attach the water bottle containing 4-6cm H²O to the exhaust? Thank you.

Unfortunately, it is not possible to finely adjust flow or pressure on the TOPO. What I can tell you is that I have it set so that the little ball reaches the 0.4L/min mark at the maximum, I have very loosely attached the inflow tubing so that the gas is being effectively sampled (I have 95% O² supplied and technically they could be ventilated on room air anyway), and I only have tubing (approx 6-8") connected to the exhaust on the back, no water bottle. Irrespective of the settings (which will depend on how it is calibrated which can vary greatly between users), you should watch the lungs themselves to make sure you don't see atelectasis, bubbling (suggestive of piercing), or overinflation (lungs look distended like an overstretched balloon, take up majority of space in the chest cavity, and may even come out when the chest is open). I wish I could be of more help. There are certainly other ventilators, for example from Columbia instruments and the Harvard apparatus Inspira (I just sent mine back again for the second time since I bought it 3.5 years ago because it is quite finicky), but they are often much more expensive and still require significant user acclimation. I'm sorry I can't be more helpful but if you have more questions, I will do my best. Good luck!

For establishing a rat MI model, I'v tried veeeeeeeery much and as guidline I used your article and two other articles in this site, but in all cases my rats have dead a few minutes after surgury, I checkd my ventilator and all other of my instruments but I couldn't find any reason. I'm so confused, because at the first of the surgery the heart rate of animal is normal but it gradually becoms slow and slower!! is there any significant diference between rat and mouse?would you help me by awaring me if I have any pitfall?

I don't personally do rat surgery now but my grad student is doing it for her post-doctoral work. She gave me the information below. Hope it helps!
suture for chest and muscle layers: Ethicon 4-0 PLAIN GUT (H8²1)
suture for the actual ligation: Ethicon Proline 7-0 (8648)
suture for skin layer: Ethicon 5-0 Coated vicryl Plus Antibacterial (VCP433)
For intubation, we have a styrofoam board with a rubber band, similar to what you
did for the mice... put the rubber band on its teeth and "stand" the rat up. Use
laryngoscope (but i think ours is actually an otoscope- welch allen, model ²1700,
which i can't find on their site) to insert a guide wire... you'll have to "shove" the
scope down pretty far before you see the vocal folds opening and closing. aim
between them. then tubing over the wire, remove the wire... connect to ventilator.
Ventilator is set at volume, 3.08cc, and 64bpm. We keep the weight set at 500g,
even though no rat weighs 500g. The settings are kept the same for any size rat we
do (most of our rats are ²50-300g, sprague dawleys). i keep the rat under 3%
isoflurane + oxygen the whole time.
Lay rat on its right side. i roll up some gauze to put under the chest to elevate it a
bit... easier to see.
Use thumb to feel for heartbeat (just under armpit). i make a small incision, maybe
a centimeter or a little more between ² ribs, parallel to the ribs). use rib spreaders.
get rid of pericardium, look for LAD- comes down from atrial appendage area.
white- to light pink in color. Ligate with 7-0. insert catheter behind a rib so it
dŒsn't slide out... pull out the needle part. close chest and muscle each with the
plain gut... then skin with ethicon 5-0. Attach a 1²mL syringe to catheter. pull back
all the way, then remove the syringe/cath. Give 3-4 1²mL injections of s.c. saline.
Wait for rat to breath on its own before taking off ventilator. Could take an hour or
more. Make sure isoflurane is off. Give buprenex for pain if required.

Thanks for your teaching. I have learned your steps to overcome almost all problems during the coronary artery ligation, including permenant ligation or ischemia/reperfusion. Now I have two problems cannot be resolved in the two mouse model:
1. myocardial I/R: ²4 hours after reperfusion, I intubed the mouse and reoccluded the LAD again. I injected 1% Evan's blue (in PBS) into the aorta root, or the catheter cannulated in carotid artery or from the right ventricle. I just can get pale blue in the myocardium, or only showing in the surrounding of the transversely myocardium. There is no clear borderline to distinguish the AAR and normal muscle. Would you please help me?
². Chronic heart failure (permenant ligation): Four weeks after ligation, I put the whole heart into the heart matrix to get the ²mm transverse slice. The muscle wall is thin and cannot have good slice. Have you any idea or procedure ?
Many thanks for your patience.

1) I recommend that you try the method of Bohl, et al described in AJP Heart and Circ ²97:H²054-²058, ²009. We have found this method to be superior in terms of clear demarcation of the perfused tissue vs AAR.
²) We do not use the matrix because our Zn-based fixative dŒs not make the tissue firm enough. And as you have noted, the 4 wk post-MI heart is particularly delicate because of the thin wall. For all histology, I use a scalpel and cut the heart into 4 transverse sections by hand. I hold the heart gently with forceps to avoid concave sections. Depending the on the resolution you need, you could try to freeze the heart briefly in saran wrap (for 15-²0min) or put it into agar. Many others have reported using these techniques so you should be able to find more detail.
Good luck!

Dear doctor Jitka Virag
I'm a master student from China ,Now do some work about MI model ,I flow your article .when I do the surgery" suture underneath the left anterior descending coronary artery (along the long axis of the heart) "but do not see any change for the myocardial and myocardial turn it white I do not know what the problem is
,I do it for a long time ,It must be very kind of you to tell me How to determine the ligation position How can I improve? Thank you very much
best wish
yours sincersly
Dezhong yang